Proteases are a class of enzymes that catalytically hydrolyze peptide bonds. Their primary chemical sequence and unique three-dimensional structure determine their activity and specificity. Depending on the active site composition, proteases are classified into major groups including aspartic, metallo-, thiol-, and serine proteases. The role of proteases in physiological processes is widely recognized. Not only are they involved in such functions as digestion, blood coagulation and fibrinolysis (Lottenberg, R.; Christensen, U.; Jackson, C. M.; Coleman, P. L., Assay of Coagulation Proteases Using Peptide Chromogenic and Fluorogenic Substrates; Methods in Enzymology 1981, 80, 341-361), but also in ovulation, tumorigenicity, immune response, and viral and bacterial infection, etc. (Livingston, D. C.; Brocklehurst, J. R.; Cannon, J. F.; Leytus, S. P.; Wehrly, J. A.; Peltz, S. W.; Peltz, G. A.; Mangel, W. F., Synthesis and characterization of a new fluorogenic active-site titrant of serine protease; Biochem. 1981, 20, 4298-4306). For example, retroviruses such as HIV are known to encode a protease which functions to process precursor proteins at specific cleavage sites. These cleavages occur during the virion assembly and are required for the maturation of infectious virus particles. Thus, inhibition of these proteases has become an important target for the design of antiviral agents, including those for AIDS.
In addition, public awareness of antibiotic resistant bacteria strains and food-borne illnesses is increasing. The management of microbial risks in healthcare, cosmetics, food and beverage industries is a serious health and safety issue. Bacterial testing is an integral part of managing microbial risks. The ability of many bacteria to produce proteases is a widely used criterion for identification and characterization of certain pathogenic species.
Sensitive and quantitative enzyme assays are required for the discovery and understanding of biological functions of proteases, the diagnosis of physiological disorders and the development of therapeutical drugs. A variety of techniques have been used to measure protease activities, including enzyme-linked immunosorbent assays (ELISA), high performance liquid chromatography (HPLC), protein immunoblot analysis, and thin layer electrophoretic analysis. However, these methods usually required multiple steps and multiple reagents, and are slow and expensive to operate. They are sometimes impractical for applications such as high-throughput screening of pharmaceutical drugs, e.g., protease inhibitors.
Fluorogenic substrates are molecules that change from nonfluorescent to highly fluorescent upon enzymatic hydrolysis. They are widely used as molecular probes for studies and tests of viral and bacterial proteases, nucleases, saccharidases, phosphatases and kinases (Manafi, M.; Kneifel, W.; Bascomb, S., Fluorogenic and Chromogenic Substrates used in Bacterial Diagnostics; Microbiological Reviews 1991, 55, 335-348). The fluorescence can be readily observed under UV illumination, by a fluorescent microscope, in a 96-well plate reader, or in a flow cytometer.
Several fluorogenic protease substrates are available commercially. Examples include EnzCheck™ kits (Molecular Probes, Inc., Eugene, Oreg.), which use highly-quenched casein substrates bearing 4,4-difluoro-4-borata-3a-azonia-4a-aza-s-indacene fluorophores sold under the trade name BODIPY (Molecular Probes, Eugene, Oreg.). When cleaved, fluorescent BODIPY-peptides are released. Typically, a two-fold increase in fluorescence intensity, at 530 nm, is observed for trypsin concentrations of up to 500 ng/mL. A fluorogenic substrate for HIV protease is available (Molecular Probes, Inc.) that includes the HIV protease cleavage site and, on one side thereof, the fluorophore 5-((2-aminoethyl)amino)naphthalene-1-sulfonic acid (EDANS) and on the other side thereof, the acceptor chromophore 4-(5-dimethylaminophenyl)azobenzene sulfonic acid (DABCYL). As described in European Patent Application No. 428,000, EDANS fluorescence is quenched by the DABCYL chromophore through intramolecular resonance energy transfer, a process requiring that the donor and acceptor be separated by no more than 100 Angstroms along the peptide chain. The fluorophore is excited by radiation at 340 nm and fluoresces at 490 nm, which can be obscured by absorbance or fluorescence of the bacterial culture medium.
U.S. Pat. No. 4,314,936 describes an enzyme assay substrate comprising an uninterrupted peptide chain to which is attached a fluorescent group in one part of the peptide and a fluorescence quenching group in another part. Cleavage of the chain at any point between the two liberates the fluorophore for detection and quantification. Specific amino acid sequences are prepared for specific enzymes. Fluorophores include eosinyl-, rhodaminyl-, and fluroesceinyl-type dyes as well as EDANS-type moieties. Quenching species (which are not dyes or fluorophores) include nitrosated aromatic compounds such as nitrophenyl, nitrobenzyloxycarbonyl, nitrobenzoyl, etc.
PCT Patent Application No. WO 95/03429 describes an immunoassay procedure wherein a fluorogenic tracer comprises a short antigen-mimicking peptide labeled with both a fluorescent energy transfer donor and a fluorescent energy acceptor. When free in solution, the tracer exhibits very little fluorescence due to intramolecular dye dimerization (quenching); when bound to an antibody of the native antigen, fluorescence is considerably increased due to dissociation of the molecular dimer brought about by conformational changes in the tracer peptide. Representative fluorescent energy dyes that form intramolecular dimers include the fluorescein family, such as fluorescein, tetramethyl rhodamine, rhodamine B, and Texas Red. Thus, the application describes fluorescence enhancement upon binding rather than fluorescence upon cleaving, and fluorescence quenching relies on a combination of energy transfer and dye dimerization.
U.S. Pat. Nos. 4,822,746 and 5,254,477 describe quantitative and qualitative analysis of analytes that relies upon the interaction of a fluorophore with a chromophoric light absorbing compound or with a second (light absorbing) fluorophore. Quenching occurs via both radiative and non-radiative energy transfer by the fluorophore when in the excited state, rather than by dye dimerization. By this means, the method of '746 is able to produce only a 10-20% increase in fluorescence in the presence of an analyte.
U.S. Pat. No. 5,605,809 describes peptides useful for protease detection, the peptides having a fluorophore conjugated to each end and folded such that the fluorophores exhibit quenching via intramolecular energy transfer. When cleaved by a target protease, the fluorophores are released from close proximity and the resulting signal is detected and quantified. FIGS. 2A and 2B of '809 show that, at most, an 9-fold increase in fluorescence is seen on cleavage of the substrate. A large number of peptides, ranging in size from 2 to about 8, preferably 2 to about 6 amino acids in length, is described. The fluorescent indicators absorb and emit light in the visible region (400-700 nm).
Fluorescence dye quenching commonly takes place by a number of mechanisms, including energy transfer and dye dimerization. In both cases, when a molecule comprising a fluorescent dye donor and an acceptor (wherein acceptor may or may not be a fluorescent dye in the case of energy transfer) linked by a chain X is excited by input of energy, typically by irradiation with a specific wavelength of light, energy is transferred from the donor dye to the acceptor rather than being dissipated by fluorescence. Energy Transfer, also referred to as Förster-type dipole-dipole interaction, generally takes place over a longer distance between donor and acceptor (on the order of 100 Angstroms). See, for example, L. Stryer et al., Energy Transfer: Spectroscopic Ruler; Biochemistry, 1967, 58, 719-726. Dye Dimerization or Dye Stacking, on the other hand, occurs when two or more fluorescence molecules are separated by a short-enough distance for their planar aromatic rings to interact to form aggregates such as dimers and trimers. The absorbance spectra of dyes in a dimer-or stacked state are substantially different from those of the same dye in energy transfer pairs. Dye dimer absorption spectra show characteristic decrease in the principal absorption peak as dye concentration increases, while showing a characteristic increase in the shoulder peak. This phenomenon is commonly referred to as “band splitting.” See, for example, K. K. Rohatgi and G. S. Singhal, J. Phys. Chem., 1966, 70, 1695-1701. See also, FIG. 2 (infra). Concentration increases can be accomplished either by increasing the amount of dye in a unit volume, or by physically locating two (or more) dye molecules closely together on a linker molecule, such as a peptide or other small molecule. Dimerization or stacking takes place through the formation of ground-state complexes (i.e., through close physical contact), whereas energy transfer interactions occur through space. Because of this, these spectral changes are not seen for dyes that interact by energy transfer mechanisms.